Understanding Lipase-Deep Eutectic Solvent Interactions Towards Biocatalytic Esterification

Abstract

Deep eutectic solvents (DESs) have shown promise as a medium for extracting polar volatile fatty acids (VFAs) and in situ esterification of the extracted molecules using lipases. This solvent enhanced biocatalysis process can potentially streamline VFA separation from fermentation broth by integrating conversion and extraction steps. Two commercial lipases from Aspergillus oryzae (AoL) and Candida rugosa (CrL) were evaluated in reaction systems containing hydrophilic or hydrophobic DESs using a newly optimized lipase assay. The optimal pH for both lipases was around 5.0, with a slight reduction in activity at pH 8.0 and a significant inhibition at pH 2.0. The impact of DES concentration on lipase activity varied depending on the specific DES–lipase pairs. Most hydrophilic DESs show good compatibility with the tested lipases. Specifically for choline chloride/ethylene glycol (1:2) and choline chloride/levulinic acid (1:2), taking into account the influence of pH, CrL activity increased with DES concentration. However, the hydrophobic DES thymol/2,6-dimethoxyphenol (1:2) demonstrated enhanced inhibitory effects on both lipases. Docking simulation helped explain the ligand–protein interactions but showed limited capability in predicting the compatibility of specific DES–lipase pairs due to its constraints in simulating flexible protein structures and the complex interactions between DES components and water.

1. Introduction

Volatile fatty acids (VFAs) can be produced from organic materials, such as agricultural residues, food waste, or lignocellulosic biomass, under anaerobic conditions through acidogenic fermentation processes [1]. However, their accumulation can inhibit fermentation microorganisms, limiting VFA yields. To address this challenge, in situ extraction of VFAs has gained significant research interest as a strategy to alleviate toxicity and improve fermentation efficiency [2,3]. In addition to traditional organic solvents such as hexane and octane and ionic liquids such as trioctylamine and [P666,14][Phos], deep eutectic solvents (DESs) integrated with membrane separation have demonstrated effective VFA extraction from complex aqueous mixtures [4]. Beyond their role in extraction, certain biocompatible DESs have shown promise as reaction media for enzyme-catalyzed esterification of VFAs, offering a sustainable route to produce valuable chemicals such as solvents, flavors, and fragrances [5]. These studies highlight the potential for developing a continuous, low-energy system for VFA recovery and conversion, offering a promising solution for sustainable bioproduct synthesis [4].

Specifically, commercially available lipases from Aspergillus oryzae (AoL) and Candida rugosa (CrL) have demonstrated improved aqueous esterification selectivity when immobilized within a confined hydrophobic space [6]. Furthermore, optimizing DES components—particularly those derived from natural products such as sugars, alcohols, amino acids, and lipids—can enhance its biocompatibility with enzymes and microorganisms. By combining these advancements, we can explore the potential of using these lipases for VFA esterification in DESs, leveraging their improved performance in tailored environments. If proven feasible, this approach could streamline separation processes by integrating VFA conversion and extraction into a more efficient one-pot system.

Nevertheless, unlike most acid substrates with lower polarity that have demonstrated applicability in lipase-catalyzed esterification reactions in the presence of DESs [7], VFAs, being more polar acid molecules, have received limited attention in such systems. Notably, the hydrolysis of VFA molecules in a mixture of aqueous fermentate and DES can influence the surrounding pH, potentially impacting reaction performance. For example, previous studies have demonstrated the influence of pH levels and water content in both pure and aqueous solutions of DESs on the stability and activity of lipases [8]. Moreover, lipase activity is also affected by DESs; while its components may lead to the inactivation or denaturation of enzymes, there are also reports of improved lipase activity due to DES. These varying impacts are commonly attributed to alterations in the secondary structure of enzymes, or the presence of abundant hydrogen bonding in DESs, which can change the affinity between substrates and enzymes [9]. For instance, Nian et al. [10] demonstrated that Candida Antarctica lipase B (CALB) could be activated through the promotion of H-bonding interactions between the substrate, such as lauric acid [11], and the DES (Choline Chloride/Glycerol) within the acyl-binding pocket. Therefore, it is essential to investigate the effects of pH, DES components, and their concentrations on lipase activity in this complex system containing aqueous VFA in fermentate and DESs.

Lipase activity is commonly assessed using the catalytic hydrolysis of p-nitrophenyl esters as a model reaction [12]. The hydrolysis activity of lipases in aqueous DES solutions, determined using this method, is widely acknowledged as an indicator of biocompatibility between a specific DES and enzyme pair [7]. However, no comparative study has been conducted to evaluate the impact of hydrophobic DESs on lipase activity using this model reaction. When considering different lipases, their conformational characteristics are known to influence their interfacial activation properties. A distinct structural feature observed in most lipase highlights the importance of solvent polarity in their activation [8]. Specifically, a polypeptide flap or lid has been found to occlude the active site, which contains a catalytic triad [13]. The hydrophobic side of the flap, predominantly composed of aliphatic side chains, faces the active site, while the hydrophilic side is oriented opposite and interacts with the protein surface [13]. Given this well-characterized structural attribute, docking simulations may be employed to potentially assess the compatibility of specific lipases with DESs. To explore the feasibility of developing a predictive model for such interactions, AoL and CrL serve as ideal candidates, as they have been extensively studied and confirmed to exhibit this common structural characteristic. Moreover, these lipases are commercially available and hold potential for practical applications in the context of this study. For instance, the lipase AoL, also known as Lipolase^® [14], is a widely recognized commercial enzyme in detergent that is originally derived from Thermomyces lanuginose [15]. However, despite their potential as predictive method candidates and their commercial significance, limited research has focused on the activity of these lipases in the presence of DES.

To address these problems, in this study, we investigated the compatibility of various hydrophilic and hydrophobic DESs with lipase AoL and CrL using an optimized hydrolysis activity assay with p-nitrophenyl butyrate (p-NPB) as the substrate. Six choline chloride based hydrophilic DESs were examined in aqueous solutions at varying DES concentrations. The impact of pH values induced by DESs was also investigated. To assess the impact of hydrophobic DESs, a biphasic system employing the same model reaction was developed. In addition, docking simulations were conducted to study the interaction between lipases and the various DES components, aiming to establish a simplified rational method for predicting the effect of different DES species on lipase activity.

2. Results

2.1. Optimized Lipase Activity Assay Method on a 96-Well Plate

Introducing DESs into a lipase activity assay can cause complications due to the pH shift, imbalance of reactant and biocatalyst, and possible phase separatation when a hydrophobic DES is used. Based on the standard lipase assay, the ratio of aqueous supernatant, water, and HEPES-NaOH buffer in the neutralization step was optimized to facilitate a clear reading and enable a linear relationship with the concentration. The reaction time for each condition was optimized to ensure that the concentration of produced p-NP falls within the range of the standard curve. The neutralization reagent volume and reaction time for each condition are listed in

Table 1. Optimized reaction time and neutralization reagent volume in hydrolysis activity assay.

Buffer/DES	Reaction Time (min)	Sample (µL)	HEPES-NaOH Buffer (µL)	Water (µL)
pH 2.0	30	50	50	-
pH 5.0	15	50	50	-
pH 7.0	15	30	10	60
pH 8.0	15	20	50	30
U5	15	20	50	30
U20	15	10	90	-
U50	15	10	90	-
Gly5	15	30	10	60
Gly20	15	20	10	70
Gly50	15	10	10	80
EG5	15	30	10	60
EG20	15	20	10	70
EG50	15	20	10	70
Lev5	30	60	40	-
Lev20	90	10	90	-
Lev50	90	10	90	-
Lac5	30	50	50	-
Lac20	90	70	180	-
Oxa5	90	40	60	-
Lev5 (n)	15	50	50	-
Lev20 (n)	15	40	60	-
Lev50 (n)	15	20	80	-
Lac5 (n)	15	50	50	-
Lac20 (n)	15	40	60	-
Lac50 (n)	15	40	60	-
Oxa5 (n)	15	50	50	-
Thy/DMP	30	50	50	-

. The optimized lipase assay, which can be performed on a 96-well plate, presents a fast and efficient method for determining and comparing lipase activity in various aqueous solutions. In contrast to prior studies where a high enzyme and substrate solution content of 19% (v/v) was induced into the final reaction system [8,16], we have reduced this value to 6% to minimize the dilution of the targeted buffer/DES solutions, leading to a more precise evaluation of its effect on lipases. Additionally, this scaled-down reaction system (on a 96-well plate) enables high throughput screening of multiple conditions simultaneously and reduced reagent consumption [16]. Moreover, the inclusion of a neutralization step expands the applicability of this method, allowing the determination of acidic buffers [17]. Notably, a high concentration of HEPES buffer at pH 7.5 demonstrated optimal performance as a neutralizing agent. It effectively stains the existing p-NP in the system, while preventing the conversion of p-NPB into p-NP, which was observed when using strong alkaline agents like NaOH solution.

2.2. Lipase Activity in Hydrophilic Non-Acidic DESs

The hydrolysis activities of lipases AoL and CrL in various pH buffers and non-acidic hydrophilic DES solutions are depicted in Figure 1. The x-axis of the figure displays abbreviations of DES names, followed by their concentration in water (w/w). In this context, U, Gly, and EG represent DES ChCl/U (1:2), ChCl/Gly (1:2), and ChCl/EG (1:2), respectively. The results of lipase activities in different pH buffers emphasize the critical impact of pH in the reaction system. Both lipases exhibited increased hydrolysis activity as the pH decreased from 8.0 to 5.0.

The pH of the ChCl/Urea (1:2) solution increased from 7.2 to 7.8 as the DES concentration was raised from 5% to 50%. Comparing the lipase activities in this DES solution with those in corresponding pH buffers, it became evident that CrL could tolerate ChCl/Urea (1:2) concentrations of up to 20% without significant activity changes. However, when the concentration increased to 50%, there was a slight reduction in activity. In contrast, AoL exhibited slight inhibition in the presence of ChCl/Urea (1:2), and the degree of inhibition increased with higher DES concentrations.

For ChCl/Gly (1:2) and ChCl/EG (1:2) solutions, the pH values remained consistent across varying concentrations, ranging between 5.1 and 6.0. The activities of CrL and AoL in the ChCl/Gly (1:2) solution were slightly lower than their pH buffer counterparts, but no significant variations were observed among different concentrations. This suggests a mild inhibitory effect of ChCl/Gly (1:2) on the lipases’ activities. Similarly, there was limited inhibition observed in the ChCl/EG (1:2) solution. Interestingly, the lowest activity was found in the 5% solution, and there was an increasing trend of lipase activities as the concentration of ChCl/EG (1:2) increased. Therefore, ChCl/EG (1:2) shows potential as a lipase activity booster rather than an inhibitor. This phenomenon is supported by previous experiments demonstrating the effectiveness of EG as a co-solvent in lipase activity assays and possible promotion of H-bonding interactions between the substrate and lipase [12].

2.3. Lipase Activity in Hydrophilic Acidic DESs

Figure 2 illustrates the hydrolysis activities of lipases under low-pH conditions induced by buffer and acidic hydrophilic DESs. The x-axis of the graph was labeled following a similar convention as in Figure 1, where Lev, Lac, and Oxa represent DES ChCl/Lev (1:2), ChCl/Lac (1:2), and ChCl/Oxa (1:1), respectively. While lipase AoL was found to be completely inactive under the pH 2.0 condition in buffer, lipase CrL exhibited better tolerance to this low-pH environment. Increasing the concentration of ChCl/Lev (1:2) and ChCl/Lac (1:2) DESs resulted in a decrease in pH. The presence of both DESs at a concentration of 5% in the system did not significantly inhibit the lipases beyond the impact of reduced pH. However, increasing the concentration of these DESs to 20% resulted in a synergistic inhibition effect in addition to the inhibition caused by low pH alone. ChCl/Oxa (1:1) had the lowest pH value, with a mere 5% concentration resulting in a pH as low as 1.2, leading to complete loss of hydrolysis activity for both lipases. DES solutions containing 50% ChCl/Lac (1:2) and 20% or 50% ChCl/Oxa (1:1) were not examined due to the expected complete inhibition of lipase activity, following the observed trend. Taking together, acidic DESs show strong inhibition to lipase when compared with neutral DESs, possibly owing to both the pH and the DES constitutents.

To eliminate the influence of pH, all acidic DES solutions were neutralized to the optimal pH of 5.0 for both lipases. The lipase activity in these neutralized DES solutions was then reassessed. The results are presented in Figure 3, with the x-axis labeling method following the same convention as in Figure 2. The (n) notation denotes the neutralized DES solutions.

After adjusting the pH to a neutral level, the inhibitory effect of acidic DESs was alleviated. Interestingly, increasing the concentration of ChCl/Lev (1:2) even resulted in an enhancement of lipase CrL activity. However, the activity of AoL was slightly inhibited by both 5% and 20% ChCl/Lev (1:2) solutions, and the inhibition effect notably increased when the concentration was raised to 50%. In the case of neutralized ChCl/Lac (1:2) DES solutions, similar activities were observed for lipase CrL compared to the pH 5.0 buffer, regardless of the DES concentration. AoL showed a slight inactivation in the presence of this DES, and the inactivation effect intensified as the concentration increased to 50%. A slight reduction in lipase activities was also observed in the neutralized ChCl/Oxa (1:1) solution. The evaluation of ChCl/Oxa (1:1) solutions with concentrations equal to or higher than 20% was not conducted due to the instability of this DES at such high concentrations in water.

2.4. Lipase Activity in Hydrophobic DESs

Hydrophobic DESs are particularly effective in liquid–liquid extraction processes, where they selectively separate polar or non-polar compounds from aqueous solutions. Several studies have demonstrated that hydrophobic DESs such as menthol/lauric acid [18], Men/Dmp [4], TOAB/DA, and Thy/DMP [6] can achieve an extraction efficiency of greater than 80% for VFAs in fermentation processes. We screened the TOAB/DA and Thy/DMP hydrophobic DESs in this study. Since TOAB/DA (1:2) completely inhibited lipase activity at even the lowest DES concentrations, Thy/DMP (1:2) was employed as a representative hydrophobic DES to assess its compatibility with lipases in the biphasic reaction system. Both lipases exhibited low activities: 14.6 U/mg for CrL and 4.2 U/mg for AoL, respectively. Compared with most of the hydrophilic DESs, hydrophobic DESs appear to cause stronger inhibition. The inhibitory effect was particularly emphasized, considering the enhanced driving force resulting from increased substrate concentration in this system. The lower lipase activity could be rooted in the inhibitory effect of the hydrophobic DES components but also caused by the poor mass transfer of the two-phase system. The interface of the aqueous phase and the DES phase creates a barrier for the migration of the substrate, where the structure of the DES and hydrogen bonding influence the cross-phase movement of organic substrates by affecting the partition and kinetics in biphasic systems [19].

2.5. Docking Simulations

To better understand the interaction of DESs with the lipase, we conducted protein–ligand docking simulations. It is noted that two lipases are structurally different from each other, while both can exhibit open and closed conformations (Figure 4). Since the active site is accessible to a ligand or an inhibitor in the “open” state, when performing docking simulations, we mainly focused on the open conformation to predict the likelihood of a DES binding to the active site. A DES occupying/binding to the active site will cause a possible inhibition effect due to interference with substrate binding.

Figure 5 illustrates the top-ranking outcome from 25 runs of induced fit docking, revealing the lowest free energy of binding. Notably, the “#” symbol positioned above certain bars indicates that the ligand binds to the side chain of the active site during global docking. This suggests a greater likelihood of the ligand binding to the active site, in comparison to other regions of the lipase protein.

Based on the docking results, the hydrophobic DES group demonstrated the strongest interaction with the lipases, as indicated by its lowest free energy of binding. Global docking simulations revealed that both lipases allowed the Thy molecule and TOA cation to access their active sites. These interactions were primarily attributed to hydrophobic interactions, as supported by the results of a contact analysis conducted through the YASARA Structure software (version 23.2.25) (ESI Figure S1). The significant affinity observed between the hydrophobic DES components and the lipases may have played a role in inhibiting the hydrolysis activity of the lipases, as discussed earlier.

The hydrophilic DES groups, on the other hand, exhibited overall lower affinity towards the lipases. The common constituent of all hydrophilic DESs, choline chloride (ChCl), demonstrated a relatively higher binding energy of >−4 kcal/mol for both choline and chloride ions. The other constituents in non-acidic hydrophilic DESs, including urea (U), glycerol (Gly), and ethylene glycol (EG), displayed energies of binding to both lipases that were comparable to those observed for ChCl (also shown in the contact analysis in ESI Figure S2). The acidic hydrophilic DES components, levulinic acid (Lev), lactic acid (Lac), and oxalic acid (Oxa), exhibited slightly weaker binding energies, indicating a moderate level of affinity (also shown in the contact analysis in ESI Figure S3). Considering the diverse effects observed in the previous hydrolysis activity assays, where different hydrophilic DESs contributed to either inhibition or enhancement, it is difficult to establish a universal rule regarding the relationship between hydrophilic DES affinity and inhibitory performance towards lipases.

This outcome reveals a limitation of using docking simulations in predicting the compatibility of lipase activity with DESs. Firstly, docking simulations rely on a fixed lipase structure that is either in an open or closed conformation, which inadequately represents the true structure of lipases within a complex reaction system. The protein structure, particularly the conformational changes in the lid, is influenced by several factors, such as the polarity of hydrophilic DES components, interfacial characteristics in the biphasic system with hydrophobic DESs, and the pH value of the reaction system, all of which were not accounted for in the docking approach. Secondly, accurately predicting the effects of interactions between the two components of DES on their affinity to lipases, as well as the modifications within the DES system caused by the presence of water, remains challenging using the docking simulation approach. Our simulations only accounted for the individual interactions between each component and the lipases, failing to capture the true complexities of the system. For instance, previous studies have demonstrated that the interaction between ChCl and U that contributes to the formation of DESs also aids in restricting the diffusion of the individual U molecules to the protein core, thereby mitigating denaturation of the CALB lipase [21].

3. Discussion

Our study on the compatibility of various DESs with lipases AoL and CrL has yielded several important insights into the potential use of these systems for biocatalytic applications, particularly in the context of one-pot VFA extraction and esterification.

3.1. Lipase Activity Assay Development

The optimized lipase assay method developed in this study offers significant advantages over previous approaches. By reducing the enzyme and substrate content from 19% to 6% [8,16], we achieved a more precise evaluation of DES effects on lipase activity while minimizing reagent consumption. This high-throughput, 96-well plate format enables efficient screening of multiple conditions, which is crucial for optimizing aqueous DES–lipase systems. The inclusion of a neutralization step extends the method’s applicability to acidic buffers, addressing a common limitation in lipase activity assays.

3.2. pH Effects and Lipase Performance

Both lipases exhibited increased hydrolysis activity as pH decreased from 8.0 to 5.0, with significant inhibition at a pH of 2.0. This pH-dependent behavior aligns with previous studies on lipase activity in various media [8]. The superior performance of CrL compared to AoL in DES environments suggests that the enzyme source and structure play crucial roles in determining compatibility with non-conventional solvents [22].

After considering pH effects, most hydrophilic DESs showed limited inhibitory effects on lipase activity, indicating good overall compatibility. The varying effects of DES concentration on lipase activity highlight the complexity of these systems and the need for careful optimization in practical applications [23]. This concentration effect also implicitly reflects viscosity-related influences, though explicit rheological characterization was beyond our current scope. It is also noted that a conventional potentiometric pH meter was used to measure the pH of aqueous DES solutions, which may have limitations in these complex water-soluble organic solvent systems [24]. Future work could adapt a unified pH measurement for DES–water mixtures, but it may require specialized calibration methods [24,25]. The relative pH comparisons in this study remain valid within the scope of our work focusing on enzymatic activity trends.

The enhanced inhibitory effects of hydrophobic DES Thy/DMP on both lipases, supported by docking simulation results, provide insights into the molecular interactions at play. This strong hydrophobic interaction with the lipase’s active site may be beneficial for VFA extraction but could pose challenges for simultaneous esterification reactions [26]. Future research could explore strategies to mitigate this inhibition while maintaining extraction efficiency.

3.3. Limitations of Docking Simulations

The use of docking simulations as a simplified molecular simulation approach has limitations in capturing the flexibility of protein structures and considering the interactions between DES components and with water molecules [27]. Consequently, docking simulations proved to be inefficient in predicting the compatibility between DESs and lipases. In contrast, molecular dynamic (MD) simulations have demonstrated efficiency in determining interactions among DES components, water, and targeted analytes in extraction processes and provide an effective method for explaining extraction performance [28]. In our previous theoretical study, we found that water molecules weaken the solvation shell of the lipase protein by reducing the protein–DES hydrogen bond lifetimes [29]. This highlights the importance of investigating the H bond network between DES components and the interaction of water molecules with DES components and lipase. Therefore, in future research, we recommend developing an MD simulation system that can simulate the interaction of lipase protein molecules in a more comprehensive system and can take into consideration factors such as pH, hydrogen bonds, and the interactions between DES components and water. In doing so, a more effective inhibitory prediction system can be established for screening lipase compatible DESs.

In conclusion, this study provides a foundation for understanding lipase behavior in aqueous DES systems, particularly in the context of VFA processing. We demonstrated reduced but still substantial activity of lipases in the presence of hydrophobic DESs. The complex interplay between enzyme structure, solvent properties, and reaction conditions highlights the need for a multifaceted approach to optimizing these biocatalytic systems. By combining experimental studies with advanced computational modeling and process engineering, we can work towards developing efficient, sustainable processes for VFA valorization and other applications in green chemistry and biorefining.

4. Materials and Methods

4.1. Enzyme Purification

Lipases AoL and CrL were purchased from Sigma-Aldrich (Saint Louis, MO, USA), with the product numbers L0777 and L1754, respectively. Prior to application, CrL was dispersed in deionized (DI) water at a concentration of 1 mg/mL and subsequently passed through VWR^® syringe filters (0.45 µm, Nylon, Radnor, PA, USA) to eliminate any insoluble impurities. The filtrate was then concentrated and underwent three washes with DI water using an Amicon^® Ultra-15 centrifugal filter unit (Darmstadt, Germany) with a molecular weight cutoff (MWCO) of 10 kDa. The AoL, a liquid product, was directly diluted to a concentration of 1 mg/mL and then subjected to the same ultrafiltration protocol as the CrL. The protein concentration in the concentrated lipase solutions was determined using the Pierce™ Coomassie (Bradford) protein assay kit (Rockford, IL, USA).

4.2. DES Preparation

Five hydrophilic DESs, including ChCl/Gly (1:2), ChCl/EG (1:2), ChCl/Lev (1:2), ChCl/Lac (1:2), ChCl/Oxa (1:1), and ChCl/U (1:2), and two hydrophobic DESs, including Thy/DMP (1:2), and TOAB/DA (1:2), were examined in the current study, where ChCl, Gly, EG, Lev, Lac, Oxa, U, Thy, DMP, TOAB, and DA stands for choline chloride, glycerol, ethylene glycol, levulinic acid, lactic acid, oxalic acid, urea, thymol, 2,6-dimethoxyphenol, tetraoctylammonium bromide, and decanoic acid, respectively. To prepare the DESs, the two components were mixed according to the noted molar ratio in parenthesis and stirred at 65~80 °C for 1~2 h until forming a homogeneous liquid. The prepared DESs were cooled down to room temperature and stored in a desiccator before usage.

4.3. Enzyme Activity Measurement

A modified spectrophotometric assay employing p-NPB as the substrate was applied to determine the hydrolysis activity of lipases in a DES or buffer solution. To determine the effect of pH, lipase activity was tested in phosphate buffers (0.1 M) with pH values adjusted to 2.0, 5.0, 7.0, and 8.0. The lipase activity was also determined in a hydrophilic DES solution prepared using deionized water at varying DES concentrations (w/w) of 5%, 20%, and 50%, respectively. Additionally, to diminish the impact of reduced pH caused by acidic DESs such as ChCl/Lev (1:2), ChCl/Lac (1:2) and ChCl/Oxa (1:1), the lipase activity in these DESs was reassessed after their pH was adjusted to neutral with NaOH solution (10 M).

For each pH buffer and hydrophilic DES solution, an endpoint assay was applied, wherein a reactant mixture of 235 µL buffer or DES solution and 10 µL p-NPB solution in EG (0.2 M) was prepared and equilibrated at 40 °C for at least 30 min, followed by the addition of 5 µL purified enzyme solution to initiate the reaction. After incubating at 40 °C for a specific duration, the reaction was terminated by introducing 100 µL of Marmur solution (chloroform: isoamyl alcohol, 24:1), followed by centrifugation at 6000 rpm for 2 min at 4 °C [30]. The resulting aqueous phase was neutralized by mixing it with a specific volume of DI water and a 40% pH 7.5 HEPES-NaOH buffer [17]. Subsequently, the absorbance of the neutralized sample at 405 nm was measured using a UV spectroscopy (SpetraMax M2 plate reader, Molecular Devices, San Jose, CA, USA).

A biphasic reaction system was used to test lipase activity in the presence of hydrophobic DESs. The reaction system was composed of 120 µL of DI water, 115 µL of DES, 10 µL of substrate solution, and 5 µL of enzyme solution. These constituents were added in the same order as the monophasic system described earlier. The reaction system was rotated at a moderate speed of around 30 rpm during incubation to ensure sufficient contact between the enzymes and the hydrophobic DES phase. To terminate the reaction, the mixture was directly centrifuged at 4 °C after the optimized reaction time. The aqueous supernatant was neutralized and subjected to absorbance measurement using the same method as described above. It is worth noting that in the preliminary experiment with the p-nitrophenol (p-NP) product standards, a decrease in the reading was observed, possibly due to the extraction of the colorimetric p-NP compound into the DES phase. To address this problem, the concentration of the substrate solution was increased to 0.4 M, and dimethyl sulfoxide (DMSO) was used to dissolve the higher amount of p-NPB substrate [31].

To rule out the background absorbance of the buffer and DES solutions, a blank assay was conducted for each condition by substituting the enzyme solution with 5 µL DI water. A standard curve of the p-NP product was generated following the same procedure used for the blank samples, except that a series of p-NP solutions with known concentrations in the corresponding cosolvent/emulsifier (EG or DMSO) were added instead of the substrate solution. The reaction time for each condition was optimized to ensure that the concentration of produced p-NP fell within the range of the standard curve. The neutralization reagent volume and reaction time for each condition are listed in Table 1.

All experiments were run in triplicate. The hydrolysis activity was calculated using the following equation: ${\mathrm{A}}_{\mathrm{h}\mathrm{y}\mathrm{d}\mathrm{r}\mathrm{o}\mathrm{l}}={\displaystyle \frac{\left({\mathrm{c}}_{\mathrm{p}\mathrm{N}\mathrm{P}}-{\mathrm{c}}_{\mathrm{b}\mathrm{l}\mathrm{k}}\right)\times {\mathrm{V}}_{\mathrm{p}\mathrm{N}\mathrm{P}\mathrm{B}}}{\mathrm{t}\times {\mathrm{m}}_{\mathrm{p}}}}$, where A_hydrol (IU/mg) represents the specific lipase activity for the hydrolysis of p-NPB; c_p-NP (mM) is the concentration of p-NP in the enzyme-containing sample after the reaction; c_blk (mM) is the concentration of p-NP in the blank sample after reaction; V_p-NPB (mL) is the volume of the p-NPB solution; t (min) is the incubation time; and m_p (mg) is the protein weight of the applied lipase.

4.4. Docking Simulation Methodology

In order to investigate the potential interactions between DES components and lipases, we conducted docking simulations utilizing the YASARA Structure software package (version 23.2.25, YASARA Biosciences GmbH, Vienna, Austria) with the AutoDock and VINA plugin. The structures of DES components were obtained from the PubChem website in SDF files. The three-dimensional crystal structures of the target proteins were acquired from the RCSB PDB website (www.rcsb.org, 27 March 2025) and underwent a cleaning process to remove water molecules and unwanted residues by applying PyMOL. Specifically, the open conformation of both the AoL (PDB ID: 1dte) and CrL (PDB ID: 1crl) lipase was employed. Each constituent molecule or ion that forms the DES was treated as a distinct ligand, while the target lipase molecule served as the receptor. Both the ligand and receptor files underwent energy minimization using YASARA Structure before proceeding with the docking simulation [32].

Firstly, a global docking approach was employed to dock each ligand onto the rigid target protein. This step aims to examine all regions within the energy-minimized protein structure to identify the most accessible binding sites. Subsequently, induced fit docking was carried out at specific active sites (Ser 209, Glu 341, and His 449 for 1crl; Ser 146, Asp 201, and His 258 for 1dte) aiming to assess the binding potential of each ligand to the active site and determine the corresponding binding energy. During the induced fit docking, only the atoms of the side chains at the essential active site within the simulation cell were treated as flexible, while the backbone and other side chains were held fixed. For each simulation condition, 25 runs were conducted.

5. Conclusions

In this study, we have developed an efficient lipase assay method that reduces reagent consumption while exhibiting extensive applicability across different aqueous–DES systems and pH ranges. This method was used to determine the activity of two commercial lipases, CrL and AoL, in the presence of hydrophilic and hydrophobic DESs. Generally, lipase CrL exhibited higher activity and tolerance compared to AoL. The pH of the system played a crucial role in lipase performance. When the pH increased from 5.0 to 8.0, both lipases exhibited a slight reduction in activity, with AoL decreasing from 26.5 IU/mg to 17.1 IU/mg and CrL decreasing from 33.2 IU/mg to 24.3 IU/mg, respectively. However, a significant inhibition was observed at a pH of 2.0, emphasizing a marked sensitivity to acidic conditions. After accounting for pH effects, most hydrophilic DESs showed limited inhibitory effects, indicating a good compatibility with the lipases. The impact of DES concentration in water on lipase activity varied depending on the DES–lipase pair. Notably, CrL exhibited an increasing trend in activity as the DES concentration increased from 5% to 50% for DESs ChCl/EG (1:2) and ChCl/Lev (1:2). In contrast, the hydrophobic DES Thy/DMP (1:2) demonstrated enhanced inhibitory effects on both lipases, likely due to its strong hydrophobic interaction with the lipase’s active site, as evidenced by docking simulation results. While docking simulation was explored as a potential method for predicting the compatibility of specific DES–lipase pairs, its limitations, such as the inability to simulate flexible protein structures and consider the interactions between the DES components and water in the aqueous system, rendered it inefficient in explaining most experimental data or predicting their compatibility.